Upload
others
View
2
Download
0
Embed Size (px)
Citation preview
Regeneration in sponge Sycon ciliatum mimics postlarval development
Anael Soubigou1,3, Ethan G Ross2,3, Yousef Touhami3, Nathan Chrismas3, Vengamanaidu Modepalli3*
1 Sorbonne University, Faculté de Sciences et Ingénierie, Campus Pierre et Marie Curie Jussieu, Paris 75252, France 2 University of Southampton, School of Ocean and Earth Science, Southampton, SO17 1BJ, United Kingdom 3 Marine Biological Association of the UK, The Laboratory, Citadel Hill, Plymouth PL1 2PB, United Kingdom
*Corresponding author: [email protected]
Abstract
Somatic cells dissociated from an adult sponge can re-organize and develop into a functional
juvenile. However, the extent to which regeneration recapitulates embryonic developmental
signaling pathways has remained enigmatic for more than a century. To this end, we have
standardized and established a sponge Sycon ciliatum regeneration protocol to achieve
consistent regeneration in cell culture. From the morphological analysis, we demonstrated that
dissociated sponge cells follow a series of morphological events resembling embryonic and
postlarval development. Hence, we propose that sponge regeneration represents somatic
development. To support our hypothesis, we performed high-throughput sequencing on
regenerating samples and compared the data with regular embryonic and postlarval
development of Sycon ciliatum. Our comparative transcriptomic analysis illuminates that
sponge regeneration is equally as dynamic as embryogenesis. We find that sponge regeneration
is orchestrated by complex regulatory mechanisms by recruiting signaling pathways like those
utilized in embryonic development to organize into a functional juvenile. In the current study,
we lay down the basic framework to study Sycon ciliatum regeneration. Since sponges are
likely to be the first branch of extant multicellular animal and the sister lineage to nearly all
animals, we suggest that this system can be explored to study the genetic features underlying
the evolution of multicellularity and regeneration.
Introduction
The gradient of regeneration widely differs in the animal kingdom and among them, the
sponge is an exceptional organism with an extraordinary regenerative ability. Anatomical
plasticity and cell differentiation is a characteristic feature of sponges, and many sponge cells
.CC-BY-NC-ND 4.0 International licensemade available under a(which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is
The copyright holder for this preprintthis version posted May 5, 2020. ; https://doi.org/10.1101/2020.05.04.061218doi: bioRxiv preprint
are capable of transdifferentiation [1, 2]. Dissociated cells from an adult sponge have the
unique ability to reaggregate and fully reconstitute into a functional sponge [3-5]. Despite its
significance, we still lack an in-depth understanding of the molecular mechanisms and cellular
basis of sponge regeneration. Sponges are the ancient extant branch of metazoans (branching
order is still debated) [6] and understanding their cellular behavior or morphogenesis during
regeneration can provide insights into the evolution of multicellularity and regeneration.
Calcareous sponges have been extensively studied in the past centuries and analysis of their
development has significantly influenced evolutionary theory. Terms such as gastrulation and
metamorphosis were first coined in syconoid species from the classic Calcaronea studies [7].
Sycon ciliatum, a calcareous sponge, is widely explored in embryonic and regeneration studies
[8, 9].
In S. ciliatum, embryogenesis is viviparous. In brief, after fertilization the embryos
through early cleavages develop into a rhomboid shape. Subsequent cell divisions result in the
formation of a cup-shaped embryo (stomoblastula). The embryo is composed of granular
macromeres adjacent to the choanocytes and ciliated micromeres pointing into the embryonic
cavity. After inversion, the cilia are positioned on the outer surface of the larva and eventually
the ciliated amphiblastula larva migrates into the radial chamber and swims out of its parent
(Fig.1) [8-10]. The amphiblastula larva is composed of a single layer of embryonic cells:
ciliated micromeres on the anterior pole, and non-ciliated granular macromeres on the posterior
pole. During metamorphosis, the ciliated cells of the anterior half undergo an epithelial-to-
mesenchymal transition and form the inner cell mass; the choanocytes will differentiate from
these cells. The macromeres envelop the inner cell mass and gradually transform into
pinacocytes (pinacoderm). After 24 hours following settlement, the spicules (monaxons) are
produced by sclerocytes which have differentiated from the inner cell mass. A choanocyte
chamber is formed in the inner cell mass and develops into the choanoderm. An osculum is
formed at the apical end and the juvenile is elongated along the apical-basal axis rises into a
syconoid body plan [8, 10] (Fig.1).
Many studies have been dedicated to S. ciliatum regeneration to understand their
morphological signatures, starting from Wilson 1907 [3], the pioneer of this topic. Despite
their strong morphological resemblance to embryonic and postlarval development, to date, no
study has systematically compared them. Several transcriptome studies of regeneration in
cnidarians and bilaterians have demonstrated that many developmental pathways are recruited
in regeneration [11-14]. Hence we aim to study the regeneration of S. ciliatum and compare it
to embryonic and postlarval development at both morphological and transcriptional levels. We
.CC-BY-NC-ND 4.0 International licensemade available under a(which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is
The copyright holder for this preprintthis version posted May 5, 2020. ; https://doi.org/10.1101/2020.05.04.061218doi: bioRxiv preprint
first carried out S. ciliatum regeneration to acquire morphological data for comparison to
regular embryonic and postlarval development and classified the regeneration into seven key
stages. Then, we collected the regenerating aggregates at defined time points and performed
high-throughput sequencing to compare the transcriptional profile with those of normal
embryonic and postlarval development.
Results and Discussion
Sycon ciliatum regeneration morphologically resembles embryonic and postlarval development.
First, we standardized and established a S. ciliatum regeneration protocol to achieve
consistent sponge regeneration in cell culture. Dissociated cells sink to the bottom of the dishes
within 1 hr of dissociation. Among these cells, we detected choanocytes, amoeboid cells, pieces
of spicules and round cells. The majority of the cells lose their cell morphology. In 4-6 hours
post dissociation (hpd), loose multicellular aggregates of diverse shapes (~10 µm) are formed;
they are known as primmorphs. The contacts between the cells were not strong, and upon
shaking the aggregates fell apart into separated cells. Within 24 hpd, these primmorphs
significantly increased in size (~100 µm) predominantly due to the incorporation of cells from
suspension or by fusing with other aggregates (Fig.2 A-C). The primmorphs have no defined
surface epithelium; on the surface pinacocytes and amoebocytes, as well as flagella and collars
of choanocytes were noticeable and similar types of cells can be found inside the primmorphs
(Supplementary file 1-3). During the early stages the cells are actively reorganized and at 48
hpd most of these cells have lost their typical features, primarily due to transdifferentiation
(Fig.2 J-K). The primmorphs become denser and spherical and two layers of cell types can be
identified: a uniform single-cell layer of pinacocytes on the surface and loosely packed granular
cells inside the primmorphs (Fig.2 D-F) (Supplementary file 3). Earlier studies coined this
phenomenon of forming bilayer cells as redevelopment, or somatic development [5, 15].
Choanocytes on the surface were oriented with their flagella outwards and retained their
characteristic features up to 2-3 days post dissociation (dpd) and after that, they started
gradually losing their typical features (flagellum and microvilli) and becoming
indistinguishable from other cells (Fig.2 I, L). Largely, two types of primmorphs were
observed in the culture: the first one is granulated cell packed spheroid with a small bubble and
another one is with a large bubble and a small inner cell mass (termed as blow-outs by Julian
S. Huxley. [4]), an interesting morphology that resembles a blastula with blastocoel.
.CC-BY-NC-ND 4.0 International licensemade available under a(which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is
The copyright holder for this preprintthis version posted May 5, 2020. ; https://doi.org/10.1101/2020.05.04.061218doi: bioRxiv preprint
Eventually, a single continuous cavity is formed between the external single cell layer of
pinacocytes and internal cell mass (Fig.2 H). At 3 dpd, the primmorphs were entirely covered
with pinacocytes, which may have possibly formed through transdifferentiation from
choanocytes (Fig.2 J-L).
Around 5 dpd spicules start appearing from the primmorphs and simultaneously multiple
ciliated chambers start appearing inside the cell mass. Similarly to regular postlarval
development [8], the monaxons were formed first (Fig.3 A, D). Over the next 6-10 days, the
ciliated chambers expanded and became lined with choanocyte epithelium (Fig.3 G-I)
(Supplementary file 4 & 5). Gradually, these choanocyte chambers fuse into a single cavity
lined with choanocytes (also known as choanoderm). Around 16 dpd a single extended cavity
(spongocoel) with osculum forms at the apical end, and simultaneously multiple porocytes with
ostia are formed (Fig.4 A, C) (Supplementary file 6). These juveniles continue elongating
along the apical-basal axis, and long straight spicules form a crown around the osculum.
Further development can be described as an increase in the complexity of spongocoel and the
development of intermediate porocytes with ostia (Fig.4 E-J) (Supplementary file 7).
Regeneration in S. ciliatum displayed a multitude of complex morphogenesis steps,
progressing through a series of aggregation, organization, and somatic-development events.
Based on our current observations and multiple pieces of evidence from previous studies [3-
5], we classified sponge regeneration into seven stages based on the morphological and cellular
events: I) Aggregation, II) Primmorphs, III) Primmorphs with monaxons, IV) Primmorphs with
ciliated chambers, V) Choanoderm, VI) Pre-Juvenile (Ascon stage), VII) Juvenile; the details
are presented in the following table (Table 1, 3rd column). Remarkably these morphological
signatures are vastly comparable to embryonic and postlarval development of S. ciliatum (Fig.1
and Table 1, 4th column) [1, 8, 9]. To gain further insights into their morphological overlap,
we compared regeneration with both embryonic and postlarval development. The
embryogenesis of calcaronean sponges was previously well described on light and electron
microscopy levels [1, 8, 9]. The morphological comparison was performed using the data from
the published S. ciliatum embryonic development studies (Leininger, S., et al. 2014 and Eerkes-
Medrano, D. I. and S. P. Leys 2006). As demonstrated in Table 1, apart from initial stages of
primmorphs at day 1, 2 and 3, the majority of S. ciliatum regeneration steps are overlapping
with a normal S. ciliatum development directly after metamorphosis including S2, S3, S4, S5
and YS [8].
.CC-BY-NC-ND 4.0 International licensemade available under a(which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is
The copyright holder for this preprintthis version posted May 5, 2020. ; https://doi.org/10.1101/2020.05.04.061218doi: bioRxiv preprint
The gene expression of regenerating Sycon ciliatum is as dynamic as postlarval development
A long-standing question in the field of sponge regeneration is whether and to what
degree regeneration recapitulate embryonic developmental pathways. Extensive morphological
similarities between regeneration and postlarval development (PLD) have driven us to explore
the gene expression pattern of both processes. We sequenced regenerating samples at 7-time
points spanning from 1 to 24 days including dpd 1, 2, 3-4, 6-8, 10-14, 16-18 (PJUV) and 21-
24 (JUV). For postlarval development, the RNAseq datasets spanning 7-time points were
collected from previously published data (PRJEB7138 & PRJEB5970) including
amphiblastula (swimming larva), flat post larva (S1), solid post larva (S2), hollow post larvae
(S3), choanocyte chamber (S4 & S5) and asconoid juvenile (PY) stages [8]. For comparative
analysis, we processed both data sets through similar pipelines as described in the material and
methods section. To assess the global transcriptomic profile underlying postlarval development
and regeneration, we performed principal component analysis (PCA) on individual data sets.
We found that both regeneration and PLD follow similar PCA distribution and surprisingly the
transcriptional changes during regeneration are as variant as PLD (Fig.5 A&B). We also
performed hierarchical clustering (HC) on both datasets to define the major clusters and we
identified two major clusters that we classified as early and late stages (Fig.5 C&D). From
PCA and HC analysis we observed that clusters in regeneration are organized in a hierarchy
consistent with what we observed in the PLD.
We then directly compared the transcriptomic variation among regeneration and PLD
using PCA. Despite their transcriptomic dynamics, regeneration and PLD fall apart from each
other mainly at PC1 proportion. However, the profiles of regeneration and PLD began to
converge as they progress through development (Fig.6 A). This indicates that the
transcriptomic dynamics of regeneration and PLD are probably similar at advanced stages.
Similar conclusions were also drawn in HC analysis: the late stages (D10-14, PJUV & JUV)
from regeneration have clustered with S5 and PY from PLD (Fig.6 B&C), further suggesting
that the later stages of both regeneration and PLD may express genes with maximum
correlation. High variance among early time points of regenerating and developmental samples
indicates an alternative path was taken during the early stages of regeneration. As noted from
the morphology, early stages of regeneration are not strictly a morphological recapitulation of
development with potential for extensive temporal plasticity during early regeneration.
A further comparison was carried out on the significantly differentially expressed genes
(padj (FDR) < 0.01) at any time point compared to day1 for regeneration and S1 for PLD.
.CC-BY-NC-ND 4.0 International licensemade available under a(which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is
The copyright holder for this preprintthis version posted May 5, 2020. ; https://doi.org/10.1101/2020.05.04.061218doi: bioRxiv preprint
Regeneration exhibited 8937 differentially expressed genes (DEG), which is comparable to
9847 DEG in PLD, suggesting that regeneration exhibits dynamic gene expression equivalent
to PLD. Of these dynamically expressed genes, approximately 50% of genes (4463) are
commonly expressed in both data sets (Fig.6 D), suggesting that half of the PLD genes were
deployed during regeneration. As regeneration employs 50% of PLD genes, we intended to
compare the expression profiles of these commonly differentially expressed genes. We
subjected the candidates to Fuzzy c-means clustering [16] to group the genes based on the
expression profile (Fig.7 A&B). The clustered genes were compared between regeneration and
postlarval development. We found that the majority of the regeneration gene clusters exhibited
significant overlap with specific PLD clusters (Fig.7 C&D). We conclude that regeneration
partly resembles PLD by displaying similar gene expression profiles.
Overall, we find that late stages from both regeneration and PLD have a high correlation,
thus the major variation in these data is among the early stages. Since the regeneration is not
starting from a fertilized single-cell embryo or amphiblastula larva, early regeneration stages
do not necessarily need to follow a similar pattern of gene expression profile to normal
embryonic and PLD development. Despite its rudimentary embryonic and swimming larval
morphology, the early primmorphs were not close to normal swimming larva in gene
expression profile, however the subsequent stages do resemble postlarval development (Fig.6).
The early primmorphs are unique, as observed from morphological analysis, and multiple
cellular events at this stage are expected to have unique gene expression.
Analyzing S. ciliatum regeneration gene expression
To the best of our knowledge, sponge regeneration from dissociated somatic cells has
not been examined at the transcriptome level. We analyzed the regeneration gene expression
data to identify developmental and multicellular regulatory genes. We performed a GO-term
enrichment analysis to reveal gene network acting during regeneration. The analysis revealed
that a set of transcription factors tightly controls the regeneration process. A broad range of
eumetazoan developmental regulatory genes were expressed during regeneration, gene families
such as Wnt, Tgfb, Fzd, and Smad were differentially expressed (Fig.8 A). Surprisingly, the
majority of listed genes were previously identified in S. ciliatum during embryonic and
postlarval development [8, 17-19]. From the pathway enrichment analysis we identified Wnt,
TGF-beta, Notch, Hedgehog, FGF and EGF signaling pathways, and the majority of core
pathway components were detected (Fig.8 A, Supplementary file 8). Each of these signaling
.CC-BY-NC-ND 4.0 International licensemade available under a(which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is
The copyright holder for this preprintthis version posted May 5, 2020. ; https://doi.org/10.1101/2020.05.04.061218doi: bioRxiv preprint
pathways triggers several downstream signaling cascades regulating cell behavior such as cell-
cell and cell-ECM interactions [20]. Along with other biological processes, apoptosis plays an
important role during regeneration [21, 22]. Apoptosis mediated regeneration permits cell
elimination and reorganizes cell composition. Major cell death or apoptotic associated
pathways including Wnt signaling, oxidative stress response and p38 MAPK pathway were
expressed during S. ciliatum regeneration (Supplementary file 8), revealing the importance of
apoptosis in remodeling the primmorphs to initiate re-development.
After tissue dissociation, the initial cellular source constitutes a heterogeneous pool of
adult somatic cells. During the initial regeneration phase (day 1 to 3) the dissociated cells re-
aggregate and organize into a morphologically distinguishable structure (Fig.2). During the
development of multicellular organisms, cells change their position extensively and this
process is governed by cell-adhesion and motility molecules. Similarly, the sponge
regeneration is guided through a series of cellular events including initial cell aggregation and
followed by cell organization. Hence, we analyzed the genes associated with cell adhesion and
cell mobility molecules. Using GO term analyses of DEG, we identified members associated
with cell-adhesion gene families, including those that encode cell-surface receptors,
cytoplasmic linkers, and extracellular-matrix proteins (Fig.8 B). Expression of these cell-cell
and cell-ECM adhesions signify highly integrated networks crucial for the individual cells to
adhere and organize into a multicellular structure [23]. Cadherins mediate cell-cell adhesion
broadly regulating morphological aspects during development [24, 25]. Several cadherin
pathway associated genes were significantly differentially expressed (Fig.8 B). Along with
cadherins we also identified integrin pathway associated genes. Cell migration is a process that
is highly dependent on adhesion and junction molecules such as integrins [26, 27]. The co-
expression of cadherin, integrins and other down-stream scaffolding and adaptor proteins such
as RhoGTPases, tyrosine kinases, and phosphatases, suggests that together these molecules
contribute to initiating the early events of regeneration. Integrins mediate contact between cells
and ECM in many organisms by binding to ECM proteins [28-32]. Three major extracellular
matrix structural proteins fibronectin, collagen and laminin displayed differential expression
(Fig.8 B), demonstrating the importance of ECM molecules during regeneration. In summary,
our analysis indicates that initial phases of regeneration, including cell aggregation and
organization, are guided by a set of cell-cell and cell-ECM adhesion associated molecules.
.CC-BY-NC-ND 4.0 International licensemade available under a(which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is
The copyright holder for this preprintthis version posted May 5, 2020. ; https://doi.org/10.1101/2020.05.04.061218doi: bioRxiv preprint
Lessons to learn on the origin of multicellularity over the sponge regeneration model.
The factors that have driven the emergence of multicellular animals from their unicellular
ancestors have yet to be addressed. Several studies on unicellular relatives of animals including
choanoflagellates, filastereans, and ichthyosporeans have significantly contributed to
understanding how unicellular organisms transform into multicellular structures such as
colonial choanoflagellate [33-35]. This suggests similar transitions may have taken place in the
animal ancestors, which eventually evolved into a stable and functional multicellular animal
[36, 37]. The current hypothesis on the origin of multicellular animals generally agrees that
choanoflagellates share a common ancestor with animals (Fig.9). This idea is based on the
molecular phylogeny and morphological similarity between choanoflagellates and sponge
choanocytes (Fig.9 A-C) [38]. Further, the choanoflagellate colonies partially resemble the
choanocyte chambers of the sponge [37, 39, 40]. In addition to their unicellular sister lineages,
we also need to investigate other close relatives of the first multicellular animals.
Studying the early regeneration events in sponge, including the signalling molecules
triggering the aggregation of cells, can provide a window into understanding the transition from
unicellular to multicellular life forms. Recent studies on multiple unicellular holozoans
revealed several genes related to multicellularity including cell differentiation, cell-cell, and
cell-ECM adhesion (Fig.9 D) [41-44], suggesting these factors have evolved before the origin
of multicellular animals. Indeed, in our current analysis on S. ciliatum regeneration gene
expression, we found that several cell signalling and adhesion protein families including
cadherin, integrins, and ECM molecules were highly expressed within 24 hrs from dissociation
(Fig.8 B). The main focus of the current study is to test the homology between regeneration
and embryonic development, hence sampling was prioritized based on morphology with the
first sample collected 24 hr following dissociation. To gain an in-depth understanding of the
initial signalling cascades triggering the aggregation a tight sampling during the initial
aggregation stages would be beneficial. Future studies in this direction will provide further
insights into the evolutionary mechanisms underlying the origin of multicellular animals.
Conclusions
Sponge regeneration is a powerful system to understand the cellular and molecular mechanisms
governing animal regeneration in general. This is crucial for understanding the evolution of
multicellularity. We compared regeneration and embryonic development in S. ciliatum by
applying both morphological and RNA-seq analyses to reveal the correlation among these
.CC-BY-NC-ND 4.0 International licensemade available under a(which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is
The copyright holder for this preprintthis version posted May 5, 2020. ; https://doi.org/10.1101/2020.05.04.061218doi: bioRxiv preprint
phenomenons. Here we demonstrated that S. ciliatum regeneration is governed by core
components of the Wnt, Tgfβ, Notch and Hedgehog signaling pathways, which are known to
guide various aspects of embryonic development and morphogenesis in extant eumetazoans.
Furthermore, we have identified major eumetazoan cell-adhesion gene families, including
those that encode cell-surface receptors, cytoplasmic linkers, and extracellular-matrix proteins.
We found that approximately 50% of genes differentially expressed during postlarval
development are also expressed in regeneration. The S. ciliatum dissociated cells thus deploy
a similar set of genes as their embryonic and larval development to assist regeneration in cell
culture. This has important implications for using the S. ciliatum regeneration system to model
the evolution of multicellularity. In the current study our sampling was aimed to dissect the
early cellular events in regeneration, an additional high throughput studies over multiple time
points can address deeper questions such as somatic cell trajectory during regeneration.
Methods
Animal collection and cell culture
Adult S. ciliatum specimens were collected in the vicinity of Plymouth, Devon, UK. All
specimens were at the stage of the post reproduction growth and did not contain any
reproductive elements. Prior to experimentation, animals were maintained in the aquarium with
filtered seawater (FSW) no longer than 2-4 hr. Each S. ciliatum was cut into small pieces by
using a scalpel and micro scissors, then mechanically passed through 40 µm nylon mesh
(Fisherbrand™) into a petri dish (35 mm). The dissociated cells were suspended in FSW and
pipetted multiple times to allow maximum dissociation. The dissociated cells from each
individual were distributed into six-well plates. The number of cells in the plates ranged from
1.2×106 -1.5×107. Cell cultures were maintained at 15°C in dark, every 24 hr the culture
medium was exchanged with FSW. The cell culture was observed daily, and bright-field
images were collected. Initially, when attempting to follow published protocols [3-5], we were
unsuccessful with the S. ciliatum regeneration, the majority of the cell aggregates were unable
to progress to functional juveniles. Hence, we modified the protocol by fractionating the
dissociated cells through centrifugation. We followed similar instructions until cell
dissociation, and then we centrifuged the dissociated cell at 1.2 rpm for 2 min. The supernatant
was collected into a new 1.5 ml tube and re-centrifuged at 2.5 rpm for 2 min. The cell pellets
from both steps were suspended in FSW and distributed individually in 6 well plates. The cells
pelleted at 2.5 rpm have a high regeneration ability in comparison to the cells pelleted at 1.2
.CC-BY-NC-ND 4.0 International licensemade available under a(which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is
The copyright holder for this preprintthis version posted May 5, 2020. ; https://doi.org/10.1101/2020.05.04.061218doi: bioRxiv preprint
rpm. In the current study, we used the aggregates collected from 2.5 rpm wells. Apart from
standardizing the protocol, the regeneration experiment was performed 7 times independently
and 8 to 12 specimens were used in each experiment. For morphological analysis, the samples
were collected on days post dissociation 1, 2, 3, 4, 5, 6, 8, 10, 12, 14, 18, 21 and 24. 10-20
regenerative structures were collected per replicate.
Whole-mount immunofluorescence
The specimens were fixed for 1 hr at 4°C on a roller with 0.05% glutaraldehyde and 4%
paraformaldehyde (PFA) in PBS buffer. After fixation, the samples were washed 5 times with
PBST (1× PBS, 0.05% (vol/vol) Tween-20) for 5 min. For long term storage, the specimens
were dehydrated in a series of ethanol dilutions and stored in 70% ethanol at 4°C. For whole-
mount immunofluorescence, the samples were permeabilized in Dent's fixative (80% methanol,
20% DMSO) for 15 minutes and followed by rehydration through series of methanol dilutions
(70%, 50% and 25% methanol) for 5 minutes per dilution. The samples were washed with
PBST for 3 x 10 min, then blocked in 5% BSA in PBST for 1 hr at RT. Primary antibody (1:500
dilution, mouse Anti-α-Tubulin Cat # T9026, Sigma-Aldrich) incubation was performed in a
blocking solution (1% BSA in PBST) for 24-36 hr at 4°C. The samples were washed with
PBST for 5 x 5 min, after which samples were incubated with secondary antibodies (1:250
dilution; Goat anti-Mouse IgG Alexa Fluor 594 Cat # A-11032, ThermoFisher) diluted in
blocking solution for overnight at 4°C. Then, the samples were washed with PBST for 5 x 10
min and clarified following iDISCO protocol [45]. Imaging was performed on Leica TCS SP8
DLS and Leica DMi8 confocal microscopes.
Fixation of regenerating specimens for Scanning Electron Microscopy (SEM)
The specimens were fixed overnight at 4°C by 2.5% glutaraldehyde in PBS buffer, then rinsed
thrice with PBS buffer for 10 min. The specimens were postfixed in 1% osmium tetroxide in
PBS buffer at room temperature. After post-fixation, the specimens were dehydrated in a series
of ethanol dilutions until 100% ethanol and absolute ethanol was gradually exchanged with
hexamethyldisilazane (HMDS). After the final 100% HMDS solution step, the specimens were
left in a fume hood overnight to allow HMDS to evaporate. After overnight drying, the samples
were mounted on aluminum stabs and gold coated. SEM imaging was done on JEOL 6610 LV
microscope.
.CC-BY-NC-ND 4.0 International licensemade available under a(which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is
The copyright holder for this preprintthis version posted May 5, 2020. ; https://doi.org/10.1101/2020.05.04.061218doi: bioRxiv preprint
RNA sequencing and differential gene expression
The regeneration is semi-synchronous among individuals, even though the variation is
considerably mild (generally varies ±1 day). Keeping this in mind we collected the regenerating
samples principally based on morphological signatures apart from the day 1 and 2 samples. For
RNA isolation, the early regenerating samples were collected at days 1 and 2. For later stages,
the samples were primarily collected based on the morphological state extending between day
3-4, day 6-8, day 10-14, day 16-18 (PJUV) and day 21-24 (JUV). Each sample was pooled
from a minimum of three individuals and also included samples from different batches as
detailed in table 1, 1st column. The samples were carefully collected using a pipette and excess
media was removed before snap freezing in liquid nitrogen and stored at -80°C until further
processing. Due to the sheer size of primmorphs at early stages (day 1 and 2) samples collected
from multiple biological replicates were combined to acquire an adequate amount of RNA for
sequencing. For the later stages, only 3 biological replicates were combined into one sample
during RNA isolation. Total RNA was isolated using the TRI Reagent® according to the
manufacturer’s protocol. RNA quality was assessed using Agilent RNA 6000 Nano Kit on
Agilent 2100 Bioanalyzer (Agilent, USA), and only samples with RNA integrity number (RIN)
≥ 8.0 were considered for sequencing. Sample library preparation for RNA sequencing was
accomplished using the SENSE mRNA-Seq Library Prep Kit (Lexogen GmbH). Before
sequencing, the libraries were pre-assessed by Agilent High Sensitivity DNA Kit (Agilent,
USA) and quantified using Qubit™ 1X dsDNA HS Assay Kit (Invitrogen™). The sequencing
was outsourced (GENEWIZ Illumina NovaSeq/HiSeq 2x150 bp sequencing), generating a
20 million paired-end reads per replicate. Raw data was deposited at NCBI GEO submission
GSE149471. After de-multiplexing and filtering high-quality sequencing reads, the adapter
contamination was removed by using Trimmomatic v0.36 [46]. The quality of the reads was
verified using FastQC [47]. Processed reads from each sample were mapped to the S. ciliatum
genome (indexed bowtie2 [48]) by using HISAT2 [49]. Next, the mapped reads were passed
to StringTie [50] for transcript assembly. After initial assembly, all assembled transcripts from
regeneration and development were merged by using StringTie's merge module, which merges
all the gene structures found in any of the samples. The BAM file and GFT file generated by
StringTie's merge were fed into featureCounts [51] to extract the reads counts per transcript.
Differential expression analyses were performed using DESeq2 (Galaxy Version 2.11.40.6)
[52]. Gene models that did not have >10 counts in at least 25% of the samples were excluded.
The principal component analysis (PCA), hierarchical clustering (HC), and heat maps were
.CC-BY-NC-ND 4.0 International licensemade available under a(which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is
The copyright holder for this preprintthis version posted May 5, 2020. ; https://doi.org/10.1101/2020.05.04.061218doi: bioRxiv preprint
generated using the R package in R-studio (Version 1.2.5019). For functional annotation, the
transcripts were extracted from the GTF annotation file by gffread utility and the LongOrfs
were acquired using TransDecoder. We used blastp [53] with the default curated gathering
threshold to predict the protein orthologues against the Uniprot database. The Gene Ontology
(GO) term enrichment was performed using gene annotation tools including PANTHER™
Classification System [54] and DAVID [55].
Acknowledgments
The authors would like to thank John Bishop and Christine Wood for their support in acquiring
S. ciliatum specimens. We thank Glenn Harper and the team at the Plymouth Electron
Microscopy Lab for their assistance during the imaging. We thank Ro Allen for his help with
R statistics. This work was supported by the Anne Warner endowed Fellowship through the
Marine Biological Association of the UK.
.CC-BY-NC-ND 4.0 International licensemade available under a(which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is
The copyright holder for this preprintthis version posted May 5, 2020. ; https://doi.org/10.1101/2020.05.04.061218doi: bioRxiv preprint
Figure 1: Schematic representation of embryonic and postlarval development stages in
Sycon ciliatum: depicted based on Leininger, S., et al. 2014, Eerkes-Medrano, D. I. and S.
P. Leys 2006. From left to right: oocyte, cleavage, pre-inversion, post-inversion, amphiblastula
(swimming larva), flat post larva, solid post larva, hollow post larvae, and choanocyte chamber
and asconoid juvenile.
.CC-BY-NC-ND 4.0 International licensemade available under a(which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is
The copyright holder for this preprintthis version posted May 5, 2020. ; https://doi.org/10.1101/2020.05.04.061218doi: bioRxiv preprint
Figure 2: Morphological and cellular events during the primmorphs stage. The
regeneration proceeds with the aggregation of dissociated cells and transforming the loosely
connected external cells to an intact single layer of pinacocytes. (A-C) Initial multicellular
aggregates in the culture (24 hpd), loosely aggregated primmorphs, the primmorphs have no
defined surface epithelium and made of diverse cell types. (D-F) A general view of primmorphs
in the culture after 2 dpd, (E & F) an intact single layer of cells surrounding primmorphs,
separating from inner cell mass, similar to blastocyst. (G & H) The external ameboid or
cuboidal cells transformed into flat pinacocytes, the external cavity (*) resembles the
extraembryonic cavity with a single layer of flat cells. (E, F, H, I) Fusing the external cells
surrounding the primmorphs, accompanied by the transdifferentiation of the choanocytes and
other external ameboid cells. (J-L) choanocytes transdifferentiation into pinacocytes: (J)
.CC-BY-NC-ND 4.0 International licensemade available under a(which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is
The copyright holder for this preprintthis version posted May 5, 2020. ; https://doi.org/10.1101/2020.05.04.061218doi: bioRxiv preprint
choanocytes with intact flagella and microvilli, (K & L) gradually these morphological features
were lost through transdifferentiation and forms falt later of pinacocytes. Images are taken with
(A, D, G) bright field, (B, E, H) confocal, (C, F, I) SEM microscope. P: pinacocyte; C:
choanocytes; F: flagella; M: microvilli; (*): external cavity.
.CC-BY-NC-ND 4.0 International licensemade available under a(which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is
The copyright holder for this preprintthis version posted May 5, 2020. ; https://doi.org/10.1101/2020.05.04.061218doi: bioRxiv preprint
Figure 3: Specule formation during regeneration. (A-F) general view of primmorphs
developing spicules (4-10 dpd). (D & G) Primmorphs developing multiple chambers
surrounded by ciliated cells, simultaneously the spicules start appearing as monaxons. (G-I)
expansion and integration of choanocyte chambers and eventually developing into choanoderm
with a single layer of choanocytes surrounding the atrium. (F & I) note that ostium and osculum
are not formed yet. Images are taken with (A, B, C) bright field, (D, E, F) SEM microscope,
(G, H, I) confocal. C: choanocytes; S: sclerocytes; P: pinacocyte; Mo: monaxons. (*): external
cavity.
.CC-BY-NC-ND 4.0 International licensemade available under a(which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is
The copyright holder for this preprintthis version posted May 5, 2020. ; https://doi.org/10.1101/2020.05.04.061218doi: bioRxiv preprint
.CC-BY-NC-ND 4.0 International licensemade available under a(which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is
The copyright holder for this preprintthis version posted May 5, 2020. ; https://doi.org/10.1101/2020.05.04.061218doi: bioRxiv preprint
Figure 4: Developing a functional juvenile. (A, B) General view of pre-juvenile and juvenile
stages. (C-J) Osculum opening at apical end and multiple porocytes form ostia. (C) The
appearance of ostium and osculum is synchronized. (G) Ostia in the regenerated juvenile, (H)
a view of ostia from inside, ostia is surrounded by choanocytes. O: Ostia; PO: porocytes; C:
choanocytes; S: sclerocytes ; P: pinacocyte; Os: osculum; A: archaeocytes or amoebocytes.
.CC-BY-NC-ND 4.0 International licensemade available under a(which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is
The copyright holder for this preprintthis version posted May 5, 2020. ; https://doi.org/10.1101/2020.05.04.061218doi: bioRxiv preprint
Table 1: Summary of the morphological and cellular events during regeneration in
S.ciliatum. The table details various regeneration steps and their overlaps with postlarval
development. 4th column (postlarval development) depicts the morphological events at
postlarval development of S. ciliatum. 1st column details number of samples collected for RNA-
sequencing at each stage of regeneration, N=number of pooled regenerating structures per well,
n=number of pooled regenerating structures per replicate, R=number of replicates per stage.
.CC-BY-NC-ND 4.0 International licensemade available under a(which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is
The copyright holder for this preprintthis version posted May 5, 2020. ; https://doi.org/10.1101/2020.05.04.061218doi: bioRxiv preprint
Figure 5: A global overview of postlarval and regeneration transcriptomes. (A) Principal
component analysis (PCA) of postlarval datasets, samples at SL, S1, S2, S3, S4, S5, and YS
stages. (B) PCA of regeneration dataset, samples at dpd 1, 2, 3-4, 6-8, 10-14, 16-18 (PJUV)
and 21-24 (JUV). (C & D) hierarchically clustering (HC) on postlarval development (C) and
regeneration (D) datasets revealed two major clusters that we classified as early and late stages.
This analysis confirmed that clusters in regeneration are organized in a hierarchy consistent
with what we observed in the PLD.
.CC-BY-NC-ND 4.0 International licensemade available under a(which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is
The copyright holder for this preprintthis version posted May 5, 2020. ; https://doi.org/10.1101/2020.05.04.061218doi: bioRxiv preprint
Figure 6: Global transcriptomic comparison of the regeneration and PLD samples. (A)
PCA of larval development versus regeneration datasets. Regeneration exhibits similar
transcriptomic variation as embryogenesis. The regeneration samples collected at D10-14,
PJUV and JUV stages are close to the S5 and PY stages of PLD. (B) Hierarchical clustering,
the dendrogram of both datasets obtained with pvclust is shown with the bootstrap values on
each node. The dendrogram shows a significant correlation among late stages of regeneration
(D10-14, PJUV and JUV) with S5 and PY stages of PLD. (C) Comparison of differentially
expressed genes between PLD and regeneration datasets, similarity matrix for significantly
differentially expressed genes (padj (FDR) < 0.01 for any timepoint comparison against Day 1
for regeneration and S1 for PLD). The yellow box highlighting the similarity between late
stages of regeneration (D10-14, PJUV and JUV) with S5 and PY stages of PLD. (D) The
number of differentially expressed genes in regeneration is highly comparable to PLD and
~50% of genes are commonly expressed.
.CC-BY-NC-ND 4.0 International licensemade available under a(which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is
The copyright holder for this preprintthis version posted May 5, 2020. ; https://doi.org/10.1101/2020.05.04.061218doi: bioRxiv preprint
Figure 7: Regeneration gene clusters overlapping with PLD gene clusters. A total of 4468
commonly expressed genes between regeneration (A) and PLD (B) are subjected to Fuzzy c-
.CC-BY-NC-ND 4.0 International licensemade available under a(which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is
The copyright holder for this preprintthis version posted May 5, 2020. ; https://doi.org/10.1101/2020.05.04.061218doi: bioRxiv preprint
means clustering. Based on their gene expression profile we generated 7 clusters in both
datasets. (C) Hierarchical clustering. The dendrogram of all gene expression clusters from
regeneration and PLD. Majority of regeneration gene modules clustered with PLD gene
modules. (D) Cluster overlap table of regeneration versus PLD gene expression clusters
produced via Pearson correlation analysis. Bold text indicates the best cluster overlap and their
matching gene expression profiles are presented on the right side of the table.
.CC-BY-NC-ND 4.0 International licensemade available under a(which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is
The copyright holder for this preprintthis version posted May 5, 2020. ; https://doi.org/10.1101/2020.05.04.061218doi: bioRxiv preprint
Figure 8: Gene expression across S. ciliatum regeneration. Heatmap of significantly DEG.
(A) A heatmap displaying a selected list of transcriptional factors and Wnt and Tgfb pathways
genes. The majority of listed genes are well known to associate with a range of eumetazoan
developmental regulatory pathways and were previously identified in Sycon ciliatum
embryonic development. (B) A heatmap showing relative expression of ECM and cell adhesion
and motility (integrin, cadherin, and scaffold) genes, a broad range of genes started expressing
.CC-BY-NC-ND 4.0 International licensemade available under a(which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is
The copyright holder for this preprintthis version posted May 5, 2020. ; https://doi.org/10.1101/2020.05.04.061218doi: bioRxiv preprint
within 24 hpd, possibly involved in cell-aggregation, adhesion, and motility during
regeneration. Heatmap of scaled expression (row z-score) of integrins with significantly
different expression (padj (FDR) < 0.01 for any timepoint comparison against Day 1).
.CC-BY-NC-ND 4.0 International licensemade available under a(which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is
The copyright holder for this preprintthis version posted May 5, 2020. ; https://doi.org/10.1101/2020.05.04.061218doi: bioRxiv preprint
Figure 9: The origin of animal multicellularity. (A-C) Morphological similarities between
choanocytes and choanoflagellates. (A) S. ciliatum choanocytes among dissociated cells; (B)
choanocyte chambers of sponge S. ciliatum; (C) unicellular and multicellular (colonial) forms
of choanoflagellates. (D) Phylogenetic tree presenting animals and closely related unicellular
holozoan lineages. The position of Ctenophora and Porifera is indicated as a polytomy due to
current uncertainty on their evolutionary origin. Animal ECM and cell-adhesion family
proteins found in unicellular holozoan lineages; empty boxes indicate absence.
.CC-BY-NC-ND 4.0 International licensemade available under a(which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is
The copyright holder for this preprintthis version posted May 5, 2020. ; https://doi.org/10.1101/2020.05.04.061218doi: bioRxiv preprint
Supplementary File 1: 4D and Z-stack of imaging of primmorphs after 24 hrs post
dissociation.
Supplementary File 2: 4D and Z-stack of imaging of primmorphs 2 days post dissociation.
Supplementary File 3: 4D and Z-stack of imaging of primmorphs 3 days post dissociation.
Supplementary File 4: 4D and Z-stack of imaging of structures with ciliated chambers 8
days post dissociation.
Supplementary File 5: 4D and Z-stack of imaging of regenerating structure at 12 days
post dissociation. Defined choanoderm with a single layer of choanocytes surrounding the
atrium. Note that ostium and osculum are not formed yet.
Supplementary File 6: 4D and Z-stack of imaging of pre-juvenile 18 days post
dissociation. The ostium and osculum are formed.
Supplementary File 7: 4D and Z-stack of imaging of juvenile 24 days post dissociation.
.CC-BY-NC-ND 4.0 International licensemade available under a(which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is
The copyright holder for this preprintthis version posted May 5, 2020. ; https://doi.org/10.1101/2020.05.04.061218doi: bioRxiv preprint
Supplementary File 8: Differentially expressed genes associated with cell death. (A)
Heatmap shows the relative expression of genes associated with cell death (apoptosis signaling
pathway, P38 MAPK, and response to oxidative stress). The selected list of genes is
significantly DE across the course of S. ciliatum regeneration. Note the large changes in gene
expression at Day 1 and the gradual decrease in the expression later on.
.CC-BY-NC-ND 4.0 International licensemade available under a(which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is
The copyright holder for this preprintthis version posted May 5, 2020. ; https://doi.org/10.1101/2020.05.04.061218doi: bioRxiv preprint
Reference
1. Ereskovsky, A.V., Development of Sponges from the Class Calcarea Bowerbank, 1864, in The Comparative Embryology of Sponges. 2010, Springer Netherlands: Dordrecht. p. 3-36.
2. Adamska, M., Differentiation and Transdifferentiation of Sponge Cells, in Marine Organisms as Model Systems in Biology and Medicine, M. Kloc and J.Z. Kubiak, Editors. 2018, Springer International Publishing: Cham. p. 229-253.
3. Wilson, H.V., On some phenomena of coalescence and regeneration in sponges. Journal of Experimental Zoology, 1907. 5(2): p. 245-258.
4. HUXLEY, J.S., Memoirs: Further Studies on Restitution-bodies and free Tissue-culture in Sycon. Quarterly Journal of Microscopical Science, 1921. s2-65(258): p. 293-322.
5. Korotkova, G., Comparative morphological investigations of development of sponges from dissociated cells. Transactions of Leningrad Society of Naturalists, 1972.
6. Brunet, T. and N. King, The Origin of Animal Multicellularity and Cell Differentiation. Developmental Cell, 2017. 43(2): p. 124-140.
7. E, H., Die Gastrae Theorie, die phylogenetische Classifi cation des Thierreichs und die Homologie der Keimblatter. Jena Zeitsch Naturwiss, 1874: p. 8:1–55.
8. Leininger, S., et al., Developmental gene expression provides clues to relationships between sponge and eumetazoan body plans. Nat Commun, 2014. 5: p. 3905.
9. Eerkes-Medrano, D.I. and S.P. Leys, Ultrastructure and embryonic development of a syconoid calcareous sponge. Invertebrate Biology, 2006. 125(3): p. 177-194.
10. Franzen, W., Oogenesis and larval development of Scypha ciliata (Porifera, Calcarea). Zoomorphology, 1988. 107(6): p. 349-357.
11. Onai, T., et al., XTsh3 is an essential enhancing factor of canonical Wnt signaling in Xenopus axial determination. The EMBO Journal, 2007. 26(9): p. 2350-2360.
12. Amiel, A.R., et al., Characterization of Morphological and Cellular Events Underlying Oral Regeneration in the Sea Anemone, Nematostella vectensis. International journal of molecular sciences, 2015. 16(12): p. 28449-28471.
13. Bryant, D.M., et al., A Tissue-Mapped Axolotl De Novo Transcriptome Enables Identification of Limb Regeneration Factors. Cell Rep, 2017. 18(3): p. 762-776.
14. Burton, P.M. and J.R. Finnerty, Conserved and novel gene expression between regeneration and asexual fission in Nematostella vectensis. Development Genes and Evolution, 2009. 219(2): p. 79-87.
15. Korotkova, G.P., Regeneration and somatic embryogenesis in sponges. 1970. 16. Kumar, L. and M. E Futschik, Mfuzz: a software package for soft clustering of
microarray data. Bioinformation, 2007. 2(1): p. 5-7. 17. Fortunato, S.A.V., et al., Calcisponges have a ParaHox gene and dynamic expression
of dispersed NK homeobox genes. Nature, 2014. 514(7524): p. 620-623. 18. Fortunato, S.A.V., S. Leininger, and M. Adamska, Evolution of the Pax-Six-Eya-Dach
network: the calcisponge case study. EvoDevo, 2014. 5(1): p. 23. 19. Fortunato, S., et al., Genome-wide analysis of the sox family in the calcareous sponge
Sycon ciliatum: multiple genes with unique expression patterns. EvoDevo, 2012. 3(1): p. 14.
20. Sanz-Ezquerro, J.J., A.E. Munsterberg, and S. Stricker, Editorial: Signaling Pathways in Embryonic Development. Front Cell Dev Biol, 2017. 5: p. 76.
21. Tseng, A.-S., et al., Apoptosis is required during early stages of tail regeneration in Xenopus laevis. Developmental Biology, 2007. 301(1): p. 62-69.
.CC-BY-NC-ND 4.0 International licensemade available under a(which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is
The copyright holder for this preprintthis version posted May 5, 2020. ; https://doi.org/10.1101/2020.05.04.061218doi: bioRxiv preprint
22. Pérez-Garijo, A. and H. Steller, Spreading the word: non-autonomous effects of apoptosis during development, regeneration and disease. Development, 2015. 142(19): p. 3253-3262.
23. Weber, G.F., M.A. Bjerke, and D.W. DeSimone, Integrins and cadherins join forces to form adhesive networks. J Cell Sci, 2011. 124(Pt 8): p. 1183-93.
24. Peyriéras, N., et al., Uvomorulin: a nonintegral membrane protein of early mouse embryo. Proceedings of the National Academy of Sciences, 1983. 80(20): p. 6274-6277.
25. Nichols, S.A., et al., Early evolution of animal cell signaling and adhesion genes. Proceedings of the National Academy of Sciences, 2006. 103(33): p. 12451-12456.
26. Friedl, P., Prespecification and plasticity: shifting mechanisms of cell migration. Current Opinion in Cell Biology, 2004. 16(1): p. 14-23.
27. Friedl, P., K.S. Zanker, and E.B. Brocker, Cell migration strategies in 3-D extracellular matrix: differences in morphology, cell matrix interactions, and integrin function. Microsc Res Tech, 1998. 43(5): p. 369-78.
28. Campbell, I.D. and M.J. Humphries, Integrin Structure, Activation, and Interactions. Cold Spring Harbor Perspectives in Biology, 2011. 3(3).
29. Humphries, J.D., A. Byron, and M.J. Humphries, Integrin ligands at a glance. Journal of Cell Science, 2006. 119(19): p. 3901-3903.
30. Theocharis, A.D., et al., Extracellular matrix structure. Advanced Drug Delivery Reviews, 2016. 97: p. 4-27.
31. Seebeck, F., et al., Integrins are required for tissue organization and restriction of neurogenesis in regenerating planarians. Development, 2017. 144(5): p. 795-807.
32. Daley, W.P., S.B. Peters, and M. Larsen, Extracellular matrix dynamics in development and regenerative medicine. Journal of Cell Science, 2008. 121(3): p. 255-264.
33. King, N., et al., The genome of the choanoflagellate Monosiga brevicollis and the origin of metazoans. Nature, 2008. 451(7180): p. 783-788.
34. Ratcliff, W.C., et al., Experimental evolution of multicellularity. Proceedings of the National Academy of Sciences, 2012. 109(5): p. 1595-1600.
35. Shalchian-Tabrizi, K., et al., Multigene Phylogeny of Choanozoa and the Origin of Animals. PLOS ONE, 2008. 3(5): p. e2098.
36. Sebé-Pedrós, A., B.M. Degnan, and I. Ruiz-Trillo, The origin of Metazoa: a unicellular perspective. Nature Reviews Genetics, 2017. 18(8): p. 498-512.
37. Adamska, M., Sponges as models to study emergence of complex animals. Current Opinion in Genetics & Development, 2016. 39: p. 21-28.
38. Laundon, D., et al., The architecture of cell differentiation in choanoflagellates and sponge choanocytes. PLOS Biology, 2019. 17(4): p. e3000226.
39. Nielsen, C., Six major steps in animal evolution: are we derived sponge larvae? Evolution & Development, 2008. 10(2): p. 241-257.
40. Brunet, T., et al., Light-regulated collective contractility in a multicellular choanoflagellate. Science, 2019. 366(6463): p. 326-334.
41. Hehenberger, E., et al., Novel Predators Reshape Holozoan Phylogeny and Reveal the Presence of a Two-Component Signaling System in the Ancestor of Animals. Curr Biol, 2017. 27(13): p. 2043-2050 e6.
42. Sogabe, S., et al., Pluripotency and the origin of animal multicellularity. Nature, 2019. 570(7762): p. 519-522.
43. Suga, H., et al., The Capsaspora genome reveals a complex unicellular prehistory of animals. Nature communications, 2013. 4: p. 2325-2325.
.CC-BY-NC-ND 4.0 International licensemade available under a(which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is
The copyright holder for this preprintthis version posted May 5, 2020. ; https://doi.org/10.1101/2020.05.04.061218doi: bioRxiv preprint
44. King, N., C.T. Hittinger, and S.B. Carroll, Evolution of Key Cell Signaling and Adhesion Protein Families Predates Animal Origins. Science, 2003. 301(5631): p. 361-363.
45. Renier, N., et al., iDISCO: a simple, rapid method to immunolabel large tissue samples for volume imaging. Cell, 2014. 159(4): p. 896-910.
46. Bolger, A.M., M. Lohse, and B. Usadel, Trimmomatic: a flexible trimmer for Illumina sequence data. Bioinformatics, 2014. 30(15): p. 2114-2120.
47. S, A., FastQC: a quality control tool for high throughput sequence data. Available online at: http://www.bioinformatics.babraham.ac.uk/projects/fastqc. 2010.
48. Langmead, B. and S.L. Salzberg, Fast gapped-read alignment with Bowtie 2. Nature methods, 2012. 9(4): p. 357-359.
49. Kim, D., et al., Graph-based genome alignment and genotyping with HISAT2 and HISAT-genotype. Nature Biotechnology, 2019. 37(8): p. 907-915.
50. Pertea, M., et al., StringTie enables improved reconstruction of a transcriptome from RNA-seq reads. Nature Biotechnology, 2015. 33(3): p. 290-295.
51. Liao, Y., G.K. Smyth, and W. Shi, featureCounts: an efficient general purpose program for assigning sequence reads to genomic features. Bioinformatics, 2013. 30(7): p. 923-930.
52. Love, M.I., W. Huber, and S. Anders, Moderated estimation of fold change and dispersion for RNA-seq data with DESeq2. Genome Biology, 2014. 15(12): p. 550.
53. Altschul, S.F., et al., Gapped BLAST and PSI-BLAST: a new generation of protein database search programs. Nucleic Acids Research, 1997. 25(17): p. 3389-3402.
54. Mi, H., A. Muruganujan, and P.D. Thomas, PANTHER in 2013: modeling the evolution of gene function, and other gene attributes, in the context of phylogenetic trees. Nucleic Acids Research, 2012. 41(D1): p. D377-D386.
55. Huang da, W., B.T. Sherman, and R.A. Lempicki, Bioinformatics enrichment tools: paths toward the comprehensive functional analysis of large gene lists. Nucleic Acids Res, 2009. 37(1): p. 1-13.
.CC-BY-NC-ND 4.0 International licensemade available under a(which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is
The copyright holder for this preprintthis version posted May 5, 2020. ; https://doi.org/10.1101/2020.05.04.061218doi: bioRxiv preprint